Troubleshooting Tip 2: The transfection efficiency of my mammalian cell culture is very low. How can I improve this?
The transfection efficiency of my mammalian cell culture is very low. How can I improve this?
The low magnification phase contrast (left) and fluorescence micrographs (right) above show cultured cells that have been co-transfected with GFP in order to easily identify transfected cells via fluorescence and thus assess transfection efficiency. There is a lot of variability in the optimal transfection conditions for each mammalian cell type, and a protocol that typically works well for one cell type might work poorly for another one. The most important factor in optimizing transfection efficiency is selecting the proper transfection protocol. This choice comes down to either one of the following protocols: calcium phosphate–mediated gene transfer, electroporation, liposome-mediated transfection, and nucleofection. These protocols can be used with most cell types, both for transient gene expression and stable transformation. Nucleofection is particularly suited for primary cells with limited dividing potential (Current Protocols in Neuroscience, Unit 4.32). Here we review issues that apply to all protocols, as well as ways to help optimize transfection efficiency in electroporation and liposome-mediated transfection.
Possible Causes and Solutions
I. Problems relevant to all protocols
Are my cells healthy? Is the cell plating density optimal?
1. For high-efficiency transfection it is important to always use a consistent number of healthy, proliferating cells. Efficiency is greatest when cells are kept in mid-log growth. Culture and passage cells in a consistent manner since transfection efficiency is sensitive to culture confluency. Plate cells so that they are 50-90% confluent on the day of the transfection. If a decrease in transfection efficiency is observed, work with a freshly thawed culture.
2. Handle cells with the greatest possible care at all times during the transfection procedure, e.g. when pipetting, to avoid mechanical damage. Excessive trypsinization will damage cells and thus lead to lower survival rates, whereas insufficient trypsinization will lead to aggregates, resulting in lower transfection efficiencies.
II. Electroporation-related problems
Did I kill most of my cells during the electric shock?
1. The objective is to use a current pulse that will maintain a ~40-70% cell viability after electroporation. The maximum voltage of the shock and the duration of the electric pulse (which is determined by the capacitance of the power source) are the two parameters that you can vary to optimize electroporation. Lower capacitance (i.e. allowing more charge to be stored) results in shorter pulses (for a capacitor discharge system). Start with a low capacitance of 25 μF and a high voltage of 1500 V (for 0.4-cm cuvettes), then increase or decrease the voltage to optimize transfection. If excessive cell death occurs, lower the length of the pulse by lowering the capacitance (down to 3 μF can be tried).
2. For sensitive cells try out a low-voltage/high-capacitance setting (i.e. longer pulses). Start at 250 V/960 μF and change the voltage up to 350 V or down to 100 V in steps to determine optimal settings.
Did electroporation overheat my cells?
Keeping cells on ice can improve cell viability and lead to higher transfection efficiency, especially at high power which can lead to heating. However, under low-voltage/high-capacitance conditions, cells have been found to electroporate with higher efficiency at room temperature. Try out electroporation at both temperatures to optimize conditions for a new cell line.
Should I use a low-salt or high-salt medium for electroporation?
Low-salt, high-resistance media (e.g. HEPES-buffered mannitol, phosphate-buffered sucrose) have been used to increase pulse duration and thus increase transfection efficiency. However, high salt, low-resistance buffers such as PBS, HBS, or tissue culture medium have been succesfully used for many cell types. Try out both conditions for a new cell line.
III. Liposome-mediated transfection-related problems
Are the concentrations of DNA and lipid I use optimal?
In general, increasing the concentrations of lipid and DNA improves transfection efficiency. However, high levels of either lipid or DNA are toxic and inhibitory. You should systematically explore lipid and DNA concentrations in order to identify optimal conditions. To this end you can use a matrix of DNA and lipid concentrations to transfect cells in a 24-well plate, then scale up as needed. Start with a range of 0.2-1.6 μg DNA per well and a range of 0.5 to 5 μl lipid reagent per well.
Should I include serum and antibiotics in the DNA-lipid mixture?
Serum interferes with the formation of DNA/lipid complexes, and antibiotics may cause toxicity during transfection. Prepare DNA/lipid complexes in an antibiotic-free, serum-free medium, and plate cells in an antibiotic-free medium prior to transfection.
More information on electroporation can be found in Current Protocols in Molecular Biology, Unit 9.3 and on liposome-mediated transfection in Current Protocols in Molecular Biology, Unit 9.4. For information on the optimization of transfection efficiency have a look at Current Protocols in Cell Biology, Unit 20.7.